Abstract

Singlet oxygen (1O2) is widely believed to be the major cytotoxic agent involved in photodynamic therapy (PDT). We showed recently that measurement of the weak near infrared luminescence of 1O2 is possible in cells in vitro and tissues in vivo. Here, we investigated the relationship between the integrated luminescence signal and the in vitro PDT response of AML5 leukemia cells sensitized with aminolevulinic acid-induced protoporphyrin IX (PpIX). Sensitized cell suspensions were irradiated with pulsed 523 nm laser light at average fluence rates of 10, 25, or 50 mWcm−2 and, 1O2 luminescence measurements were made throughout the treatment. Cell survival was measured with either propidium iodide-labeled flow cytometry or colony-forming assay. The PpIX concentration in the cells, the photobleaching, and the pO2 in the cell suspensions were also monitored. There were large variations in cell survival and 1O2 generation in different experiments due to different controlled treatment parameters (fluence and fluence rate) and other uncontrolled factors (PpIX synthesis and oxygenation). However, in all of the cases, cell kill correlated strongly with the cumulative 1O2 luminescence and allowed direct estimation of the 1O2 per cell required to achieve a specific level of cell kill. This study supports the validity and potential utility of 1O2 luminescence measurement as a dosimetric tool for PDT, as well as confirming the likely role of 1O2 in porphyrin-based PDT.

INTRODUCTION

In PDT,
3
an emerging treatment for cancer and other conditions
(1, 2)
, a systemically or topically administered photosensitizer is activated in situ with a light source tuned to match the absorption spectrum of the drug. This induces photochemical production of reactive oxygen species, the most important of which is widely believed to be [1O2 (1Δg)]
(3)
. Oxidative action leads to modification or destruction of various cellular and tissue components, with consequent biological and clinical response.

Clinical dosimetry of PDT is complex, due to the many treatment factors involved
(4)
. Effective photodynamic action requires the interaction of adequate amounts of light, drug and tissue oxygen. “Explicit dosimetry” strategies attempt to measure one or more of these treatment parameters to predict the treatment outcome. Other, so-called “implicit dosimetry” strategies attempt to use surrogates for biological damage, such as photosensitizer degradation (photobleaching), as an indicator of the effective treatment dose administered. Whereas these approaches have met with some success, they have proven difficult to implement due to the many factors involved and, particularly, their dynamic interdependence. Thus, for example, the rate of photobleaching depends on the oxygen concentration in the tissue that, in turn, may change during the treatment due to either photochemical depletion of oxygen and/or treatment-induced vascular response. “Direct dosimetry” attempts to circumvent these complications by measuring the important photobiological toxin, i.e., 1O2.

1O2 is believed to be generated by the following, type-II photochemical pathway
(5)
where S0, S1, and T1 are the photosensitizer ground state, first excited singlet state, and first excited triplet state, respectively, and 3O2 and 1O2 are the ground-state triplet and first excited singlet states for oxygen, respectively. 1O2 undergoes radiative decay, and the resulting emission at 1270 nm can be readily measured in solution
(6)
.

Measurement of the radiative decay of 1O2 at 1270 nm in biological media is of interest, both in general and as a possible direct dosimetry technique
(7, 8)
. Historically, this has not been technically feasible, because the high reactivity of 1O2 with biomolecules
(5)
dramatically reduces its lifetime. We showed recently for the first time that measurements of 1O2 luminescence in both cells and tissues is possible
(9)
using a new PMT with extended spectral sensitivity and rapid response. Specifically, measurements were made in both suspensions of AML5 and P388 cells, and normal Wistar rat skin and liver, both sensitized with tetrasulfonated aluminum phthalocyanine. The photosensitizer triplet-state lifetimes and the 1O2 lifetimes were also measured, and the latter were shown to be reduced from typical values of ∼3 μs in aqueous solutions to <200 ns in vitro and in vivo.

Given that 1O2 measurements in biological medium are now technically feasible, the next logical step is to determine their utility as a dose metric. That is, does the generation of 1O2 correlate with, or ideally predict for, the photobiological response? This will also in part test the hypothesis that 1O2 is the primary cytotoxin involved in PDT. This article will report on the first of a planned series of studies, using tumor cells in vitro and cell survival as the measure of PDT response.

The experiments reported below required substantial upgrades to our experimental setup compared with that used for our initial demonstration of feasibility
(9)
, because the acquisition times for reasonable signal-to-noise ratios were too long. These changes, which include a new higher-output power laser source and increased light collection efficiency of the NIR detection optics, will be described.

The specific experiments reported here investigated the relationship between cell survival and 1O2 luminescence measured from AML5 cells during PDT. The photosensitizer used was PpIX, induced endogenously by administering ALA. The main reason for this choice was that the first in vivo studies (in progress) to investigate 1O2versus PDT response will use ALA-PpIX to replicate reported studies of a strong fluence rate dependence of the response of normal skin to ALA-PDT
(10)
. In retrospect, the use of a prodrug rather than a photosensitizer directly introduced additional variability in PpIX concentration, but as will be seen, the measurement of 1O2 circumvents this variability.

In these experiments, we varied the light fluence rate (mWcm−2) between treatments, while maintaining the same total fluence (Jcm−2) and ALA incubation concentration. The curves of survival versus fluence were significantly different between individual experiments, due to both controlled and uncontrolled variations in light, photosensitizer concentration, and/or oxygenation. However, we demonstrate that, when plotted against cumulative 1O2, the survival curves collapse to a single “universal” response curve, and will discuss the implications of this for using 1O2 luminescence as a PDT dose metric. In addition, the data enable a direct estimate of the amount of 1O2 required for cell kill in vitro.

MATERIALS AND METHODS

Theory.

A full theoretical treatment of time-resolved 1O2 luminescence measurement was described previously
(9)
. Briefly, the equation for the concentration of 1O2 as a function of time after a short excitation laser pulse can be derived from Equation A as
where N is the photon fluence (photons cm−2) in the excitation pulse, σ is the photosensitizer ground state absorption cross-section (cm2), [S0] is the concentration of the photosensitizer ground state, φD is the quantum yield of 1O2, and τT and τD are the photosensitizer triplet-state lifetime and 1O2 lifetime, respectively.

The total number of photons emitted by the radiative decay of 1O2 at 1270 nm is given by
where τR is the radiative lifetime of 1O2 in the specific environment.
Equation (C)
can be integrated over time to give the total number of photons emitted after excitation with a single laser pulse as:
Hence, the total number of 1O2 luminescence photons after each pulse is proportional to the concentration of 1O2 generated by that pulse. From our previous work
(9)
, we determined that τT = 19 ± 3 μs and τD = 0.6 ± 0.4 μs for AML5 cells in suspension incubated with tetrasulfonated aluminum phthalocyanine. Whereas we are unaware of any other published measurements of τD in cells, this value for τT is comparable with the value of 23 μs reported by Aveline et al.(11)
for P388D1 and NBT-II cells sensitized with benzoporphyrin derivative and 7.7 μs found by Truscott et al.(12)
for fibroblast suspensions sensitized with hematoporphyrin derivative. As described previously, we eliminate the contribution of prompt photosensitizer fluorescence by time gating to reject the early portion of the signal (i.e., t < 2 μs), and approximate the integral (D) by counting photons in the interval between 2 and 60 μs (i.e., 3τT) after each laser pulse and subtracting background contributions.

Apparatus.

The experimental system is shown in Fig. 1
⇓
. The new light source was a diode-pumped, Q-switched, frequency-doubled Nd:YLF laser (QG-523–500; Crystalaser Inc., Reno, NV), emitting in the green at 523 nm. The pulse repetition rate was set to 3.2 kHz, and the pulse width was ∼10 ns. A bandpass filter centered at 523 nm (10 nm bandpass, OD3 blocking; Omega Optical, Brattleboro, VT) was placed in front of the laser to remove any residual 1046 nm light. The beam was expanded using a 25-mm diameter biconcave lens with 150 mm focal length (BICV-25.4–150-UV; CVI Laser Corp., Albuquerque, NM), so that the beam diameter at the sample was ∼1 cm.

Schematic of the experimental setup used for in vitro1O2 luminescence measurements.

All of the samples were held in quartz cuvettes (1 × 1 × 4 cm; NSG Precision Cells Inc., Farmingdale, NY) mounted on a hotplate-stirrer unit (Corning model PC-120; Fisher Scientific Ltd., Nepean, Ontario, Canada). This allowed the samples to be continuously stirred and maintained at a constant temperature of 37 ± 2°C during treatment, and served also to maintain the source-sample-detector geometry constant between experiments. The cuvettes were open so that the samples were exposed to room air at the top.

The near-IR light was collected from the sample using a 50 mm diameter biconvex lens with a 50-mm focal length (01-LDX-115; Melles Griot Inc., Nepean, Ontario, Canada) at 90° to the excitation beam. The sample was positioned 38 mm from the collection lens, giving an overall system numerical aperture of 0.58. A 50-mm diameter, 1000-nm long-pass uncoated silicon filter (506-SW, OD6 blocking; IR Optical Products Inc., Farmingdale, NY) was mounted in front of this lens to block out the excitation light and minimize fluorescence generated by the collection optics.

The detector was a liquid nitrogen-cooled PMT (model R5509–42; Hamamatsu Corp., Bridgewater, NJ) with an extended spectral range from 300 to 1400 nm. The operating voltage was set to −1500 V using a high voltage power supply (model PS350/5000V-25W; Stanford Research Systems Inc., Sunnyvale, CA), at which the dark current was <1 nA, resulting in negligible dark counts. The rapid rise time of the detector (3 ns) allowed the PMT to be operated in single photon counting mode. A high-speed current preamplifier (model SR445; Stanford) was used to amplify the output of the PMT and convert it to a voltage pulse. A MCS (model SR430; Stanford), triggered by a TTL output pulse from the laser, was used to integrate photon counts in 80 ns bins after each laser pulse.

The higher laser power allowed us to use average fluence rates up to 100 mWcm−2, increasing the signal 10-fold over our previous set-up. The higher numerical aperture represented an increase in detection efficiency by a factor of 1.78, whereas the larger diameter optics and irradiation spot size allowed a larger sample interrogation volume (i.e., detector field of view), which gave an additional 4–5-fold improvement in signal. Hence, the new system has ∼2 orders of magnitude improvement in signal compared with the original system
(9)
.

PDT Experiments.

Suspensions of OCI-AML5 cells
(13)
were grown in α-mem medium with 5% fetal bovine serum (Life Technologies, Inc., Rockville, MD) and then incubated with 1 mm ALA (Sigma-Aldrich Canada Ltd., Mississauga, Ontario, Canada) for 4 h. Immediately before PDT treatment and 1O2 measurements, the cells were spun at 1000 rpm for 10 min, resuspended in fresh medium, and agitated for 10 min. They were then spun down again and resuspended in fresh medium to minimize the amount of extracellular photosensitizer present in the supernatant. The samples were then rapidly transferred to the measurement cuvette at a concentration of 50 × 106 cells/ml in a 2.5-ml volume. The time between suspension of the cells at their final concentration and the beginning of PDT treatments (and 1O2 measurements) was typically 5–10 min.

On each experimental day, three sets of cell suspensions were irradiated at fluence rates of 20, 50, or 100 mWcm−2. The corresponding treatment durations were 60 min, 30 min, and 15 min. With a sample volume of 2.5 ml, and spot size of 1 cm diameter, and, given that the sample had relatively high optical scattering and was continuously stirred, the cells were thereby exposed to average fluence rates of ∼10, 25, and 50 mWcm−2 (because roughly half the suspension volume was irradiated at any given time), giving total treatment fluences of 36, 45, and 45 Jcm−2, respectively. Experiments at each fluence rate were repeated six times on different days (using a set of 3 cell suspensions per experiment). On each day, new cell samples were harvested from the same incubation batch. The rationale for selecting the different fluence rates was to change the photodynamic oxygen depletion rate in the samples. This has been reported previously as altering the cell survival for the same delivered fluence, with the highest killing corresponding to the lowest fluence rate
(14)
.

To summarize, the following PDT treatment parameters were explicitly controlled in these experiments: ALA concentration, fluence rate, total fluence, cell concentration in the cuvette, sample volume, and irradiation geometry. The following parameters were not explicitly controlled: oxygen concentration in the suspensions, the exact time between suspension of the cells at their final concentration and the beginning of treatment/measurement, the degree of cell confluence at the time of ALA incubation and so the distribution of cells in their growth cycle, and the resulting intracellular PpIX concentration.

During the light irradiation, 20 μl aliquots containing ∼1 × 106 cells were removed from the suspensions at multiple time points (typically 10) during luminescence measurements without interruption of the irradiation. These were placed in six-well plates with 2 ml fresh medium and reincubated, so that the survival fraction could be assessed.

Viability Assays.

For most experiments (n = 12), the cell survival fraction was assessed using FC-PI, in which cells with compromised membranes take up PI and stain positive for fluorescence. Cell aliquots were maintained in darkness in an incubator for 48 h after treatment, centrifuged at 1000 rpm for 10 min, and resuspended in 1 μg/ml PI (Molecular Probes Inc., Eugene, OR) in PBS for 10 min. They were then analyzed on a flow cytometer (fluorescence-activated cell sorter; Becton Dickinson, Mountain View, CA), with argon ion-laser excitation at 488 nm. Cells that exceeded a specified red fluorescence threshold due to PI uptake were counted as dead. The surviving fraction was taken as the number of unstained cells (i.e., cells below the specified red fluorescence threshold) divided by the total number of cells counted. Note that this assay does not account for treated cells that have been lost due to disintegration and, therefore, probably overestimates the surviving fraction at high doses.

For a subset of treatment conditions (n = 6), colony-forming assays were used to verify the FC-PI findings. For this, the cell aliquots were suspended in a semisolid medium (Methocult 4531 with penicillin and streptomycin added; Stemcell Technologies, Vancouver, British Columbia, Canada) at 103, 104, or 105 cells per ml by dilution of the original 20 μl aliquot. The suspension (1.5 ml) was then plated on 35-mm diameter plastic gridded dishes in duplicate, and incubated for 12 days. The resulting colonies were counted, and the surviving fraction was taken as the number of colonies (>20 cells) at each time point divided by the number of colonies at time zero (i.e., sensitized cells with no light).

1O2 Luminescence Data Collection and Analysis.

For each experiment, 3 replicate cell suspensions were used from the same cell culture, 2 of which were sensitized with ALA, the third serving as a control. Near infrared luminescence measurements were made continuously during irradiation of all 3 of the samples by selecting each of the 3 band-pass filters in turn and single-photon counting in the time interval between 2 and 60 μs after each laser pulse. The signal was summed over 65,000 laser pulses and then corrected for the system response at that wavelength. The maximum trigger rate for the MCS was 1.6 kHz, whereas the laser operated at 3.2 kHz, so that counts were collected only from every second pulse. Because we had established previously
(9, 15)
that our system is capable of measuring intracellular 1O2 luminescence from sensitized AML5 cell suspensions by identification of its distinct 1270 nm peak, only the 1240, 1270, and 1300 nm filters were used to increase the number of data collection time points. The starting filter was alternated between 1240 and 1270 nm to minimize sampling-order bias. Each 3-wavelength time point took ∼2.5 min to collect, including filter wheel motion, photon counting, and data transfer. Hence, 6, 12, and 24 such data sets were acquired for the 15, 30, and 60 min treatments, respectively. This entire luminescence data collection process was automated.

The spectra from each of the duplicate cell samples (sensitized) were corrected for background by subtracting the mean of all of the control (unsensitized) measurements for that batch, and the magnitude of the 1270 peak was calculated for each time point during irradiation, with points ∼2.5 min apart. These two values were then averaged to give the incremental increase in 1O2 signal during treatment. The cumulative 1O2 luminescence was obtained by summing these incremental values and multiplying by 3, because the system was acquiring at 1270 nm for only 1/3 of the treatment time. The cumulative 1O2 luminescence at time points where cell aliquots were removed was then calculated using linear interpolation.

PpIX Photobleaching.

To investigate the role of PpIX photobleaching in these experiments, six additional sets of cells were irradiated with either 10 (n = 2), 25 (n = 2), or 50 mWcm−2 (n = 2). Aliquots (20 μl) were removed from the treated suspensions at multiple time points during the treatments and placed in six-well plates. The intracellular PpIX fluorescence was measured as follows. The cells from each well were pelleted by centrifuging at 1500 rpm for 10 min, washed once in PBS to remove extracellular PpIX, and recentrifuged. The cell pellet was then resuspended in 1 M perchloric acid (Sigma) to lyse the cells. After incubation for 5 min the cells were sonicated for 10 min and then centrifuged at 1500 rpm for 10 min. The supernatant containing the intracellular PpIX was removed, and the PpIX fluorescence was measured using a spectrofluorimeter (PTI LS-100; Photon Technology International, Lawrenceville, NJ). For each experiment, the values were then normalized to the initial PpIX concentration before irradiation to determine the photobleaching rate.

PpIX Synthesis.

The level of confluence in the cultures from which cell samples were taken was not controlled in these experiments. This was not intended but, in retrospect, was useful in that it led to variations in the intracellular PpIX concentration between experiments. To check the range of this variation, the relative PpIX fluorescence was measured using the same spectrofluorimetric technique as above, without the intermediate washing step. This was performed in triplicate in cells incubated with 1 mm ALA for 4 h at 1/3, 2/3, and full confluence (defined as 1 × 106 cells/ml).

Triplet Oxygen Measurements.

To investigate the role of triplet (ground state) oxygen depletion, a Clarke-style electrode (pO2 Histogram; Eppendorf Inc., Hamburg, Germany) was placed directly into the cell suspensions during PDT irradiation, at the center of the cell sample (∼1 cm below the top surface) and the oxygen partial pressure in mm Hg recorded continuously. This was then converted to percentage of pO2 by dividing by the standard conversion factor of 760 mm Hg. The probe was periodically calibrated in water at known oxygen tension. Measurements were performed on 4 sets of unsensitized cell suspensions (2 at 50 mWcm−2 and 2 with no light), and 6 sets of sensitized cells (2 each at 10, 25, and 50 mWcm−2).

RESULTS

1O2 Luminescence and Cell Viability Measurements.

Fig. 2
⇓
shows a typical set of emission spectra at different time points during irradiation in sensitized and unsensitized (control) cells, whereas the insert shows the corresponding incremental 1O2 luminescence (i.e., magnitude of the 1270 nm peak after subtracting the unsensitized-sample background value) as a function of treatment time for the same single set of sensitized cells.

Near infrared luminescence spectra at different time points during treatment at 50 mWcm−2 for single sensitized (1 mm ALA, 4 h) and control cells. The inset shows the corresponding calculated incremental 1O2 for the single sensitized suspension (after background subtraction). The time points refer to the middle of acquisition of the spectra. Also note that some spectra have been removed for clarity.

This was repeated for all 12 of the experiments for which the FC-PI assay was performed, and the cumulative 1O2 luminescence as a function of treatment fluence for all of the experiments is shown in Fig. 3A⇓
. A striking feature of these data is the large variability in the curves even for the same fluence rate. This was unexpected and, as discussed below, is attributed to the uncontrolled PpIX and pO2 levels. However, underlying this, a general trend toward lower 1O2 generation at higher fluence rate can be seen, as would be expected, at least qualitatively, due to photochemical depletion of oxygen.

A, cumulative 1O2 luminescence as a function of cumulative treatment fluence for all 12 data sets for experiments where the FC-PI assay was performed. Illustrative error bars correspond to the range of the 2 sets of sensitized cells treated on the same day. The lines are simply to guide the eye. B, surviving fraction (FC-PI assay) as a function of cumulative light fluence. This includes 12 sets of sensitized cells (as in Fig. 3A⇓
), plus 3 sets of unsensitized cells. Illustrative error bars correspond to the range for the 2 replicates for each set. C, surviving fraction as a function of cumulative 1O2 measurement from the data of A and B for sensitized cells. Note that the data have been truncated at 4 × 104 cumulative 1O2 counts due to the limited dynamic range of the FC-PI assay.

Fig. 3B⇓
shows the corresponding cell survival data using the FC-PI assay for the same 12 sets of sensitized cells, as well as 3 sets of unsensitized controls. All of these curves have been normalized individually to the viability immediately before the start of irradiation, which was generally >85%. The control cells had an average final surviving fraction of 0.96 ± 0.03, indicating that the handling of the cells and the treatment light alone did not affect cell viability significantly. The FC-PI assay allows measurement of the surviving fraction to approximately 2–3 logs. Generally, the viability of the treated cells was <1% after exposure to the full treatment light. Again, note the significant variability in survival for the same fluence rate and the trend toward increased survival with higher fluence rate.

The data from Fig. 3, A and B⇓
are replotted in Fig. 3C⇓
, which shows the cell survival versus cumulative 1O2 luminescence. The large variability in 1O2 and cell survival seen between experiments in Fig. 3, A and B⇓
, respectively, due to both controlled (fluence rate) and uncontrolled (PpIX level and oxygenation) factors is greatly reduced. As discussed in more detail below, this demonstrates that the data collapse onto a single universal curve and that the 1O2 measurement is a robust dose metric under these in vitro conditions.

The analogous results using the colony-forming assay are presented in Fig. 4
⇓
. This assay has a greater dynamic range than the FC-PI technique, enabling measurements down to surviving fractions of 10−4-10−5. It also measures proliferative capacity of the cells after treatment, rather than simply loss of membrane integrity, and so is generally considered a more relevant measure of cytotoxicity. Again, the scatter in the data for 1O2versus fluence (Fig. 4A)
⇓
and for survival versus fluence (Fig. 4B)
⇓
is markedly reduced when survival is plotted versus cumulative 1O2 (Fig. 4C)
⇓
, confirming the findings with the FC-PI assay.

Six sets of data using the colony-forming assay. A, cumulative 1O2 luminescence as a function of light fluence. B, surviving fraction as a function of fluence for the same treatments. Illustrative error bars correspond to the range between duplicate sets of sensitized cells. C, surviving fraction (colony-forming assay) as a function of 1O2 luminescence replotted from A and B.

Because neither assay distinguishes necrotic from apoptotic cell death, an apoptosis assay was performed on two additional sets of cells, using a commercial Annexin V kit according to the manufacturer’s protocol (Annexin V-FITC; Oncogene Research Products, San Diego, CA). For suspensions irradiated with 45 Jcm−2 at 25 mWcm−2, up to 40% of the treated cells stained positive for early apoptosis (i.e., postphosphatidylserine inversion but premembrane permeabilization), and an additional 26% stained for late apoptosis/necrosis (i.e., postmembrane permeabilization; Ref.
16
). This was not intended to be a complete quantitative analysis but does indicate that many of the cells were undergoing apoptosis as a result of the PDT treatment. This is consistent with other studies
(17, 18)
showing apoptosis to be the dominant mechanism of cell death after ALA-PpIX PDT of various tumor cell lines. The significance of this observation for the present work is discussed below.

Finally, we comment on the shape of the 1O2 dose-response curves. We performed a statistical analysis of these data by replotting all of the results from the individual, unaveraged experiments corresponding to the averages in Fig. 3C⇓
and Fig. 4C⇓
. The reason for this is that in retrospect, as discussed below, uncontrolled variations in pO2 and PpIX levels made it technically impossible to repeat an experiment exactly on a given day. Hence, the individual experiments were treated as separate data sets and were analyzed as shown in Fig. 5, A and B⇓
. The surviving fraction appears to be exponential with 1O2 “dose,” with the exception of a single, spurious data set from the colony-forming assay experiments (shown with open symbols in Fig. 5B⇓
), which was left out of the analysis. In addition, these survival curves appear to have little or no shoulder. Hence, for both curves, a single exponential was fit by least squares analysis, forcing the y-intercept through unity.

Surviving fraction versus1O2 luminescence for all data sets comprising Fig. 3
⇓C and Fig. 4
⇓C for the (A) FC-PI assays and (B) colony-forming assays with best single-exponential fits for each. The data have been truncated to 10−2 for the FC-PI assay and to 10−3 for the colony-forming assay. For clarity, only some representative error bars show ±1SD in the data as described in the text.

For the data from the FC-PI experiments, the curve was fit only to points where survival was ≥0.01, to be reliably within the dynamic range of the assay. Systematic errors were not known for the FC-PI assay, but were estimated using the average size of the range bars calculated in Fig. 3B⇓
. For these, the single exponential curve fitted the data well, with a χ2 per degree of freedom of 1.8 for S(x) = exp(−x/3050), where x is the cumulative 1O2 luminescence photon counts. The corresponding fit for the colony-forming assays included the first 3 logs of cell kill and gave a best fit to S(x) = exp(−x/2100) (χ2/NDF = 1.8). The uncertainties in surviving fractions could be determined directly from the counting statistics for the colonies. These fits are also plotted in Fig. 5, A and B⇓
. Whereas the calculated reduced χ2 were reasonably low, it appears that the surviving fraction versus1O2 curves may not follow a simple exponential form, particularly at higher doses. This may be indicative of a resistant subpopulation of the cells.

Experimental Variability.

As noted above, significant differences were seen in the cell survival and 1O2 measurements between experiments with the same PDT treatment parameters (Fig. 3, A and B⇓
; Fig. 4, A and B⇓
). A limited set of additional experiments was performed to check possible causes for this (although, with hindsight it was useful to have had this additional variability, because it demonstrated the value of using 1O2 luminescence as a dose metric). The factors checked were the triplet oxygen concentration in the cuvette, the relative initial intracellular concentration of PpIX, and the rate of intracellular photobleaching.

Fig. 6A⇓
shows the pO2 measured in unsensitized cell suspensions in the quartz measurement cuvette as a function of time during PDT irradiation. The initial value was 14.7 ± 4.6%, which is significantly less than the air-equilibrated value of 21%. In all of the cases, the pO2 decreased with time, reaching a minimum of ∼2% after 27 ± 9 min. No significant difference in final pO2 level was observed between unsensitized suspensions where no treatment light was used and unsensitized cells irradiated at 50 mWcm−2 (data not shown). We attribute this decrease to metabolic consumption of oxygen by the cells at the relatively high concentration (50 × 106 cells per ml) that was used to obtain a strong 1O2 luminescence signal. The large variation in the initial pO2 level is likely due to differences in the time taken between preparing the cell suspension and the start of measurements: typically 5–10 min, but occasionally 20–30 min due to minor technical problems. A sample data set for sensitized cells irradiated with 25 mWcm−2 light is also shown, indicating that oxygen depletion occurred in the sensitized suspensions at a higher rate. As shown in Fig. 6B⇓
, the depletion rate increased with increasing light fluence rate: e.g., reaching a minimum pO2 level of <1% in 3 ± 2 min for 50 mWcm−2 compared with 10 ± 2 min at 10 mWcm−2. This increased rate of oxygen depletion in sensitized suspensions is likely the result of photochemical depletion in addition to the metabolic oxygen consumption observed in the control cells. It is worth noting that, comparing suspensions irradiated with 10 and 50 mWcm−2, approximately the same total fluence was required to reach the minimum oxygen level. This would be consistent with metabolic and photochemical depletion rates being significantly higher than the reperfusion rate. It is also of interest to note the recovery of pO2 at later times in the irradiation, which is likely due to reduced photochemical oxygen consumption as the photosensitizer is bleached. A similar observation was made in spheroids by Georgakoudi et al.(19)
.

pO2 in cell suspensions in the measurement cuvette. A, pO2 as a function of time after the start of measurement/irradiation in unsensitized cell suspensions and in sensitized suspensions incubated with 1 mm ALA for 4 h and irradiated with 25 mWcm−2. The values are the average for 4 control (± 1SD) and 2 sensitized (± range) experiments. B, time for the pO2 to reach a minimum value (<2% in control suspensions, <1% in sensitized suspensions) in control and sensitized suspensions at different treatment fluence rates. (The fluence rate did not affect the measurement in the control samples.)

Significant variability was also observed in the initial concentration of PpIX in the cells. Fig. 7
⇓
shows that, in general, the intracellular PpIX fluorescence measured per cell increased with the degree of confluence at the time of ALA incubation, which was not tightly controlled in these experiments. Specifically, up to a factor of 1.5 difference in PpIX synthesis was observed over the range of confluence conditions used. A similar effect was observed by Georgakoudi et al.(20)
, who found up to a 3-fold increase in PpIX synthesis with degree of confluence in adherent EMT6 cells after incubation with 1 mm ALA (over a greater confluence range than used here).

Mean intracellular PpIX fluorescence per cell as a function of the degree of confluence at the time of ALA incubation; bars, ±1 SD. For each condition, the fluorescence has been normalized to the mean of that for fully confluent suspensions. Full confluence is defined as 106 cells ml−1. Each column represents three duplicates of 100-ml cell suspensions incubated with 1 mm ALA for 4 h.

Finally, Fig. 8
⇓
shows the normalized intracellular PpIX fluorescence as a function of cumulative treatment fluence for each of the three fluence rates. Although not statistically significant, there is a trend toward decreasing bleaching rate with increasing fluence rate. This is reasonable, based on the corresponding observed oxygen depletion rates, but these differences probably did not contribute significantly to the intraexperimental variations in response and 1O2 generation.

Intracellular PpIX fluorescence as a function of cumulative fluence for the three fluence rates. In each case the data are normalized to the value at the start of irradiation. For each time point a 20-μl aliquot containing 106 cells was analyzed. Cells were sensitized with 1 mm ALA for 4 h, and all experiments were performed in triplicate. The corresponding photobleaching rates, based on fitting single exponential curves to these data (i.e., assuming 1st-order kinetics) were: 0.03 ± 0.02, 0.06 ± 0.03, and 0.06 ± 0.03 (Jcm−2)−1 at 50, 25, and 10 mWcm−2, respectively. These are comparable with the range of 0.02–0.04 (Jcm−2)−1 observed in EMT6 spheroids by Foster and Bigelow.
4

DISCUSSION

Technical Issues.

As in our earlier reports
(5, 9, 15)
, spectral discrimination of the 1O2 luminescence was achieved using narrow-band interference filters, rather than, for example, a monochromator
(21)
to maximize the signal-to-noise ratio. Because we established previously
(9)
that this system is capable of measuring intracellular 1O2 luminescence in vitro, and given the unambiguous 1270 nm peak, the use of three spectral bands to define the 1O2 signal should be reliable.

The optical parametric oscillator laser source used previously to demonstrate the feasibility of measuring 1O2 luminescence in vitro and in vivo was not suitable for delivering PDT treatments due to its low average power, low repetition rate, and extremely high peak power (that may cause transient saturation of the photosensitizer absorption). Switching to a high-repetition rate, low peak power pulsed laser that could deliver adequate irradiances in a reasonable treatment time enabled the studies reported here. In addition, the frequency-doubled Nd:YLF laser is compact and highly reliable. Its limitation is the fixed wavelength (523 nm). Although this is suitable for exciting PpIX and other porphyrins, it limits the range of photosensitizers that can be studied and also, for in vivo experiments, gives only shallow treatment depth. Hence, we are developing a dye-cell cavity that can be pumped by this laser to provide longer wavelength tunable output.

In our earlier work
(9)
, it was necessary to use very high photosensitizer concentrations and long acquisition times to obtain usable signal-to-noise ratios. Increasing the optical collection efficiency of the system, together with the new laser source, gave significantly reduced signal acquisition times and enables PDT treatment conditions (photosensitizer concentration, light fluence rate, and treatment time) similar to those used clinically.

We noted above that the 1O2 count rate could be compared between experiments if the optical collection efficiency remained constant and if the 1O2 lifetime, τD, was consistent between experiments. Hence, special care was taken to ensure the former by tightly controlling the sample placement relative to the detector and treatment light, the treatment light spot size and location on the face of the cuvette, and the concentration of the cells in suspension so that the optical properties of the sample were constant. For the latter condition, τD depends on the photosensitizer microenvironment, i.e., on its subcellular location. Because PpIX is synthesized endogenously in the mitochondria, differences in microlocalization would be due to diffusion with time. This effect should be small in these experiments, because a fixed incubation time (4 h) was used and the variation in time to prepare and measure the cell samples after incubation is small compared with this. In addition, whereas it has been shown for some photosensitizers that the intracellular distribution may change with treatment
(22, 23)
, we are not aware of evidence for this in the case of ALA-PpIX.

Furthermore, we reported earlier
(15)
that for this cell line and incubation conditions, ∼98% of the measured 1O2 luminescence signal is intracellular in origin. Hence, the entire observed 1O2 luminescence signal can be taken as contributing to cell killing.

Whereas the FC-PI assay provides a high-throughput and accurate method for determining cell survival, it has some significant drawbacks. In particular, it has a limited dynamic range (∼2 logs of cell survival), and the leveling off in viability seen in Fig. 3, B and C⇓
may not represent the true shape of the response curve. This is supported by the results with the colony-forming assay, which is known to be accurate over a larger range (>4 logs).

1O2 Luminescence as a PDT Dose Metric.

The key finding in this study is that the large variations in cell survival seen between experiments with different controlled treatment parameters (total light fluence and fluence rate) and uncontrolled factors (most significantly, intracellular PpIX concentration and pO2 in the cell suspensions) are largely eliminated when the biological response is plotted against the cumulative 1O2 generated during the treatments. This is supported quantitatively by the low χ2/NDF of the exponential fits to the curves, for both the FC-PI and colony-forming assays.

This finding leads to two important conclusions. Firstly, it supports the hypothesis that 1O2 is the main cytotoxic photoproduct involved in PDT, at least with this photosensitizer. Deviation from the “universal” survival versus the 1O2 curve, e.g., as the 3O2 is depleted as treatment progresses, would suggest other pathways, but this is not seen. This does not rule out the possibility that other pathways may be important with a greater degree of hypoxia. It also cannot be concluded that 1O2 is the actual cytotoxin. It could be, for example, that the type II pathway is excited but represents only a partial component in the photodynamic cell killing. Future experiments, using 1O2-specific quenching agents and measuring the resulting changes in cell killing and 1O2 generation, are planned to test this.

The second conclusion is that the 1O2 luminescence is a robust PDT dose metric compared with individual parameters, such as light fluence, photosensitizer concentration, or oxygenation, all of which contribute in a dynamic and interdependent way
(4)
to the PDT response. Thus, based on Fig. 3C⇓
and Fig. 4C⇓
, the cumulative 1O2 is certainly strongly correlated with the cytotoxicity. It would be even more valuable as a dose metric if the 1O2 measure proved to be predictive of the response. That is, could one vary some other treatment parameter and still predict accurately the cell survival from the cumulative 1O2 measurement? This will be a focus of future studies.

Other Observations.

The viability versus cumulative 1O2 curves (Fig. 3C⇓
and Fig. 4C⇓
) show no evidence of a shoulder, either by eye or in the fitting. This suggests that there is not a high threshold for cell killing, which is consistent with recent findings by Lilge et al.(24)
on apoptotic cell death in normal rat brain tissue treated with Photofrin-PDT or with ALA-mediated PpIX-PDT. Our Annexin V assay indicates that a substantial fraction of the AML-5 cells undergo apoptosis 4 h after PDT, so that the apparent lack of a 1O2 shoulder may be due to the onset of apoptotic death at low doses. However, more detailed studies at low doses are required to confirm this.

The slopes of the 1O2 dose response curves (Fig. 5)
⇓
show that for the FC-PI assay, an average of 3050 ± 80 1O2 photon counts corresponded to a 1/e survival fraction. For the colony-forming assay, the corresponding value was 2100 ± 120 counts. It is not surprising that these slopes are different, given that the assays test different biological endpoints and it is reasonable that the slope for the colony-forming assay should be steeper, because this includes all of the cell death mechanisms (both somatic and proliferative), including those in subsequent generations, whereas the FC-PI assay is sensitive only to mechanisms resulting in compromised cell membranes.

The slopes of these curves can also be used to estimate the amount of 1O2 required to induce a 1/e level of cell kill as follows. On the basis of the geometric light collection efficiency of the system (∼10%), the filter and lens throughputs (∼20%), the quantum efficiency of the PMT (∼1%), the limited trigger rate of the MCS (∼50%), and the mean transmission of 1270 nm light through 10 mm of water in the cuvette (∼50%), the total number of 1O2 molecules undergoing radiative decay in the detector field-of-view is estimated as 6.1 × 107 for 3050 counts (for FC-PI) and 4.2 × 107 for the 2100 counts (for colony-forming assay). On the basis of the radiative lifetime of 1O2 in water (τR = 5.55 s) and our previously published estimate of the 1O2 lifetime in cells (τD = 0.6 μs), these correspond to 5.6 × 1014 and 3.9 × 1014 molecules of 1O2 produced in the field-of-view. Given that the cell density in the cuvette was 50 × 106 cells/ml, we estimate that ∼107 cells were in the detector field-of-view at any one time. Hence, on average ∼5.6 × 107 and 3.9 × 107 molecules of 1O2 were required, per cell, to induce a surviving fraction of 1/e for the FC-PI and CFA assays, respectively.

These values cannot be compared precisely to the findings of Farrell et al.(25)
, who determined a threshold of 0.9 mm of 1O2 for necrosis of rat liver sensitized with Photofrin, equivalent to 5 × 108 molecules of 1O2 per cell (assuming a typical cell density of 109 per ml, or ∼10 μm average cell radius). This is, however, consistent with our data, because a lower concentration of 1O2 is presumably needed to produce 1/e cell kill in vitro than full tissue necrosis. Similarly, Georgakoudi et al.(19)
estimated a threshold 1O2 dose of 12.1 mm (∼7 × 109 molecules of 1O2 per cell to induce necrosis) in EMT6 spheroids sensitized with Photofrin. This was based on an assumption that all of the cells within a hypoxic core survived treatment, whereas all of the others were killed. Because we did not observe a distinct threshold dose in vitro, this is not directly comparable with our data; however, our estimate for 1/e cell kill is lower, as required for consistency. It is also worth noting that ALA-induced PpIX and Photofrin have different subcellular localizations, and, hence, the sites of action of 1O2 are likely different due to the limited range (∼50 nm) of 1O2 diffusion in biological environments
(26)
. Hence, the two photosensitizers would probably yield different in vitro1O2 dose response relationships.

As shown in Fig. 9
⇓
, with the exception of a single outlying data set irradiated at 50 mWcm−2, the incremental rate of 1O2 production during irradiation appears to reach approximately the same terminal slope of 10 ± 4 1O2 photon counts per second. This slope is equivalent to generating 2.0 ± 0.8 × 105 radiating molecules of 1O2 per second in the sensing volume or roughly 106 radiating molecules of 1O2 per second per ml. Given the 10−7 radiative efficiency, we infer that the terminal rate of photochemical oxygen consumption in the luminescence measurement volume was ∼1013 molecules per second. This is likely controlled by the rate of oxygen diffusion from the air into the cuvette, because the initial pO2 in the suspension is depleted rapidly (Fig. 6)
⇓
. Moreover, the fact that there is a terminal 1O2 slope in all of the cases implies that the majority of the inter- and intraexperimental variability in 1O2 production (and, therefore, cell death) comes from differences in the early portion of these curves, namely from the differences in initial pO2 and PpIX concentration (which result in different rate of 1O2 production).

Cumulative 1O2 luminescence as a function of treatment time for all 12 data sets for experiments where the FC-PI assay was performed.

In addition, the initial portion of the curves (i.e., the rise portion preceding the terminal slope) shown in Fig. 9
⇓
can be compared with the molecular oxygen depletion data from Fig. 6A⇓
as follows. The rate of metabolic oxygen consumption in the control (unsensitized) suspensions was ∼0.008% pO2 per second, or 0.9 μm per second. Similarly, the rate of oxygen depletion in the sensitized suspension irradiated at 25 mWcm−2, due to both metabolic and photochemical oxygen consumption, was ∼0.34 μm per second. Hence, the rate of oxygen depletion due to photochemical consumption alone was ∼0.25 μm per second. Using a similar argument to that above, the average initial slope of 1O2 luminescence for suspensions irradiated with 25 mWcm−2 shown in Fig. 9
⇓
is equivalent to the generation of 1.2 × 1014 molecules of 1O2 per ml per second, i.e., 0.2 μm per second, which is in good agreement with our directly measured photochemical 3O2 consumption rate of 0.25 μm per second.

In conclusion, the fact that 1O2 luminescence monitoring correlated strongly with cell survival, regardless of the treatment fluence, fluence rate, triplet oxygen concentration, initial PpIX concentration, and rate of PpIX photobleaching, illustrates its potential to circumvent many of the limitations of other PDT dosimetry methods
(4)
. This will be strengthened if the planned extensions of this work to in vivo models and to testing the predictive capabilities of the 1O2 measurement are successful. We also plan to test the feasibility of making 1O2 measurements during clinical PDT treatments. However, unless there is a substantial reduction in the cost and/or complexity of the instrumentation required
(27)
, 1O2 luminescence monitoring is not likely to become a widespread, routine tool in clinical or even preclinical PDT. Hence, its most important roles may be as a gold standard for other simpler dosimetry techniques and to answer critical photobiological questions.

Acknowledgments

We thank Hamamatsu Corp., Hamamatsu City, Japan, and in particular Dr. Ken Kaufmann (Hamamatsu, Bridgewater, NJ), for supplying the PMT system, and the Canadian Foundation for Innovation and the Princess Margaret Hospital Foundation for equipment support. Dr. Richard Hill, OCI, provided the AML5 cell line, as well as the Eppendorf oxygen probe. The assistance of Natalie Boruvka, Anoja Giles, Bob Kuba, Dr. Robert Weersink, and Lynn Wong is also gratefully acknowledged.

Footnotes

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Grant support: Canadian Cancer Society under a grant from the National Cancer Institute of Canada.